AMPing down DNA replication

Bacterial Fic proteins catalyze diverse posttranslational modifications, such as AMPylation, phosphorylation, and UMPylation, and are involved in regulating bacterial filament formation. Lu et al. found that DNA yield from bacterial cultures expressing plasmids encoding Fic-1 was reduced compared to DNA yield from bacteria encoding Fic-2 or Fic-3 plasmids. This suggested that Fic-1 is involved in inhibiting DNA replication. Fic-1 AMPylated the DNA topoisomerase, needed for bacterial DNA replication, GyrB, which inhibited GyrB activity and induced SOS response. Self-AMPylation of Fic-1 inhibited its activity. The authors also identified a Fic-1 antitoxin, which they named AntF. Fic-1 expression also resulted in filament formation in bacteria through a mechanism independent of the SOS response. Thus, this study identifies one molecular target for the inhibition of DNA replication by Fic-1 and suggests that additional targets mediate the effect of Fic-1 on bacterial filament formation.

Abstract

The morphology of bacterial cells is important for virulence, evasion of the host immune system, and coping with environmental stresses. The widely distributed Fic proteins (filamentation induced by cAMP) are annotated as proteins involved in cell division because of the presence of the HPFx[D/E]GN[G/K]R motif. We showed that the presence of Fic-1 from Pseudomonas fluorescens significantly reduced the yield of plasmid DNA when expressed in Escherichia coli or P. fluorescens. Fic-1 interacted with GyrB, a subunit of DNA gyrase, which is essential for bacterial DNA replication. Fic-1 catalyzed the AMPylation of GyrB at Tyr109, a residue critical for binding ATP, and exhibited auto-AMPylation activity. Mutation of the Fic-1 auto-AMPylated site greatly reduced AMPylation activity toward itself and toward GyrB. Fic-1–dependent AMPylation of GyrB triggered the SOS response, indicative of DNA replication stress or DNA damage. Fic-1 also promoted the formation of elongated cells when the SOS response was blocked. We identified an α-inhibitor protein that we named anti–Fic-1 (AntF), encoded by a gene immediately upstream of Fic-1. AntF interacted with Fic-1, inhibited the AMPylation activity of Fic-1 for GyrB in vitro, and blocked Fic-1–mediated inhibition of DNA replication in bacteria, suggesting that Fic-1 and AntF comprise a toxin-antitoxin module. Our work establishes Fic-1 as an AMPylating enzyme that targets GyrB to inhibit DNA replication and may target other proteins to regulate bacterial morphology.

INTRODUCTION

The morphology of many bacteria changes in response to environmental cues (1). One of the most frequently observed changes for some rod-shaped bacteria is the transition between a bacillary (free-living) and a filamentous form during the different phases of the life cycles (1). For example, uropathogenic Escherichia coli (UPEC), the predominant causative agent of urinary tract infection (2), forms intracellular bacterial communities after invasion of the superficial epithelial cells of the bladder, where a subpopulation develops into filamentous cells (3, 4). The generation of filamentous bacteria is believed to confer resistance to phagocytosis by neutrophils migrating into the infection site (5, 6). In mice defective in Toll-like receptor 4 (the receptor for lipopolysaccharide), UPEC no longer forms filamentous bacteria, indicating that the host innate immune system triggers the development program that leads to filamentation (3). Transition between filamentous and rod shape has also been observed in Legionella pneumophila, the causative agent of Legionnaires’ disease, in response to temperature as well as other environmental stimuli (7, 8). Filamentous L. pneumophila appears to be more resistant to phagocytosis, but whether the immune system induces this development is unknown (7). Similarly, filamentous Salmonella enterica serovar Typhimurium cannot invade host cells (9) despite inducing membrane ruffling, indicating the importance of cell morphology in bacterial virulence.

In a model organism, such as E. coli, cell filamentation is caused by incomplete cell division. The segregation of completely replicated chromosomes is accompanied by the formation of a site for cell division, where FtsZ, the guanosine 5′-triphosphate (GTP)–binding tubulin homolog, polymerizes into a ring-like structure to form the multicomponent complex that mediates cell division (10, 11). Inhibition of FtsZ activity by factors such as SulA causes incomplete cell division and thus filamentation (12). The expression of sulA is regulated by LexA, the master regulator of the “SOS” response that senses cellular stresses, such as DNA damage and DNA replication inhibition (13, 14).

Cell filamentation occurs in an E. coli mutant grown at 43°C in a medium containing cyclic adenosine monophosphate (cAMP) (15). Further studies revealed that this mutant harbored a G55R mutation in the gene fic (filamentation induced by cAMP) (16, 17). The Fic motif has a core structure of HXFX(D/E)GNGRXXR (x, any amino acid) and is present in thousands of proteins from organisms of all taxonomic orders (18–20).

The discovery that VopS, a type III effector from Vibrio parahaemolyticus, catalyzes the transfer of the AMP moiety from adenosine 5′-triphosphate (ATP) to GTPases involved in host cytoskeleton structure (for example, Rho and Rac) revived the study of Fic proteins (19). Other studies revealed that members of this protein family catalyze diverse reactions, including AMPylation (19, 21), phosphorylcholination (6, 22), UMPylation (23), and phosphorylation (24, 25). These proteins are involved in diverse cellular processes, such as bacterial virulence (6, 19, 21, 22), protein translation in prokaryotes (24, 25), neurotransmission in fly (26), and the unfolded protein response in eukaryotic cells (27, 28). The activity of some Fic proteins is reversibly regulated by specific enzymes. For example, phosphorylcholination conferred by AnkX from L. pneumophila is enzymatically reversed by Lem3, a dephosphorylcholinase (29). The L. pneumophila effector SidD de-AMPylates the Rab1 small GTPase (30), which is AMPylated by a Fic-independent mechanism (31).

Whereas Fic proteins involved in bacterial virulence appear to be constitutively active, the activity of most Fic proteins in bacteria and those in eukaryotes is regulated by a motif with a consensus sequence of (S/T)XXXEG found either within the Fic proteins themselves or in a small protein called the α-inhibitor encoded by a gene that is often adjacent to the fic gene. In bacteria, Fic proteins and their cognate α-inhibitors constitute typical toxin-antitoxin (TA) modules (32).

Whereas the functions of Fic proteins in bacterial virulence and the cell biology of eukaryotic cells have been relatively well documented, our understanding of the function of “housekeeping” Fic proteins from diverse bacteria is limited. Many of these proteins are annotated as cell division proteins, but little is known about their physiological role beyond the filamentation phenotype originally described for the FicG55RE. coli mutant (16). Here, we showed that Fic-1, a Fic protein from a pseudomonad, AMPylated and inhibited the DNA gyrase subunit B (GyrB), which inhibited bacterial DNA replication. Fic-1 also arrested bacterial growth and induced cell filamentation. Our results indicated that Fic-1 targeted GyrB to inhibit bacterial division and suggested that additional targets are important for the effects of Fic-1 on cell morphology.

RESULTS

Most of the Fic proteins are found in taxonomically diverse bacteria. Such a wide distribution suggests that they function in fundamental cellular processes. To determine the function of housekeeping Fic proteins, we examined potentially discernable phenotypes in E. coli by expressing such proteins from various bacteria. Among them, Pseudomonas fluorescens strain 2P24, which is used for biocontrol of certain plant diseases (33), encodes three predicted Fic proteins, which we designated Fic-1, Fic-2, and Fic-3. These proteins are 199, 342, and 395 residues long, respectively (fig. S1), with Fic-1 having a length similar to Fic proteins found in diverse bacteria (20) (fig. S2).

We studied the function of Fic-1, Fic-2, and Fic-3 proteins by expressing them in E. coli by individually introducing the coding sequence and an upstream sequence containing the putative promoter into a ColE1 plasmid. E. coli expressing Fic-1 from this plasmid produced significantly less plasmid DNA than E. coli expressing Fic-2 or Fic-3 (Fig. 1A). The inhibition of plasmid DNA yield depended on the Fic motif because a mutation in His135, which is critical for catalytic activity, eliminated the inhibition (Fig. 1B). Expression of Fic-1 in cells harboring other types of plasmids also significantly decreased their yield, suggesting that Fic-1 inhibits bacterial DNA replication (fig. S3). In contrast, expression of constitutively active forms of Fic-2 and Fic-3, which contain mutations in the predicted intramolecular inhibitory motif known to abolish the inhibitory effects (32), did not detectably affect plasmid yield (fig. S4).

(A) The effect of individual fic genes on the amount of plasmid DNA in E. coli. Plasmid DNA containing the indicated constructs isolated from saturated cultures of E. coli was resolved by agarose gel electrophoresis before (left panel) or after digestion with restriction enzymes Bam HI and Sal I (fic-1) or Xba I and Hind III (fic-2 and fic-3) (middle panel). In the middle panel, both bands represent the products of the digested plasmid DNA. Data are representative of three experiments. The intensity of the bands corresponding to the vector was measured from three independent experiments (right panel). ***P < 0.001. (B) The effect of mutating the Fic domain of Fic-1 on the amount of plasmid DNA in E. coli. Plasmid DNA containing wild-type fic-1 and fic-1 with a Fic domain mutation (Fic-1H135A) was isolated from saturated cultures of E. coli, subjected to restriction enzyme digestion, and separated by agarose gel electrophoresis (upper panel). Western blot analysis of isocitrate dehydrogenase (ICDH), a metabolic enzyme, served as a loading control. Data are representative of three experiments. The intensity of the bands corresponding to the vector was measured from three independent experiments (right panel). ***P < 0.001.

Expression of Fic-1 causes bacterial filamentation

To further determine the effects of Fic-1, we examined the morphology of cells expressing Fic-1. In E. coli strain BL21(DE3), transformants harboring a plasmid that directs the production of His6-tagged Fic-1 formed smaller colonies on LB medium even without the inducer isopropyl β-d-1-thiogalactopyranoside (IPTG) (fig. S5A). The Fic motif–dependent inhibition of E. coli growth by Fic-1 was also apparent in liquid LB medium. Whereas the strain harboring plasmid control or the one expressing the H135A mutant reached the stationary phase of growth within 12 hours, we observed little growth in cultures of the strain harboring the plasmid expressing wild-type Fic-1 in the first 14 hours of incubation (fig. S5B). E. coli cells expressing wild-type Fic-1 grew 8 hours after inoculation (fig. S5B). We speculate that such growth may be due to accumulation of mutations in fic-1 or in the bacterial chromosome, thus making the cells less sensitive to Fic-1, or a combination of both.

Microscopic analysis revealed that a fraction of cells expressing Fic-1 were filamentous with multiple (>4) distinct nucleoids. Shorter cells with two to three nucleoids were also present (Fig. 2A). In contrast, in samples of the strain expressing Fic-1H135A, no filamentous cells were found, which is similar to the strain containing the plasmid control (Fig. 2A).

(A) Cells from E. coli transformants expressing SUMO-tagged Fic-1 or its H135A mutant grown for 16 hours on LB agar resuspended in phosphate-buffered saline (PBS) were fixed and stained with Hoechst. (B) Cells of P. fluorescens grown in LB with glucose were diluted in ABM medium containing 0.2% arabinose and grown for 12 hours. Cells were treated for imaging as described in (A). Images acquired with a fluorescence microscope were pseudocolored with IPLab software. (C and D) Cells grown as described in (A) and (B) were processed for SDS-PAGE. Proteins were probed for Fic-1 by immunoblotting. The metabolic enzyme ICDH was probed as a loading control (lower panels). Similar results were obtained from at least three independent experiments.

We examined the function of Fic-1 in P. fluorescens strain 2P24. Deletion of fic-1 did not result in any discernable phenotype. We could not express Fic-1 in P. fluorescens using several constitutive promoters; therefore, we used the arabinose-inducible promoter. Induction of Fic-1 inhibited the growth of P. fluorescens, and this inhibition required a functional Fic domain (fig. S5C). Cell growth in cultures expressing Fic-1 resumed 32 hours after inoculation (fig. S5C), which we again speculate may arise through mutations in either fic-1 or its targets that rendered them no longer sensitive to its activity. A portion of cells expressing Fic-1, but not the enzymatically inactive mutant, became filamentous (Fig. 2B), suggesting that Fic-1 targets pathways conserved between E. coli and P. fluorescens. Both the wild-type and the H135A mutant were detected in E. coli (Fig. 2C); in P. fluorescens, the amount of wild-type Fic-1 was lower than that of the H135A mutant (Fig. 2D). Together, these results indicated that overexpression of Fic-1 regulates one or more cellular processes relevant to DNA replication, bacterial growth, or both.

Fic-1 interacts with GyrB, a subunit of DNA gyrase

We sought to determine the mechanism of action of Fic-1 by identifying its cellular target. Because Fic-1 inhibited plasmid replication, we predicted that one or more proteins involved in DNA replication may be involved. To this end, we examined the potential interactions of Fic-1 with each of the 77 E. coli proteins known to participate in DNA replication (table S1). Among these, GyrB, a subunit of DNA gyrase (type II topoisomerase) specifically interacted with Fic-1 in a bacterial two-hybrid assay (34). We detected significant β-galactosidase activity only in the E. coli strain coexpressing the appropriate domains of two-hybrid assay protein Cya fused to Fic-1 and GyrB, respectively (Fig. 3A). When recombinant GyrB was incubated with beads coated with Fic-1, the Fic-1H135A mutant, or bovine serum albumin (BSA), beads coated with Fic-1 or Fic-1H135A retained more GyrB (Fig. 3B). Thus, by interacting with GyrB, Fic-1 may regulate bacterial DNA replication.

(A) Fic-1 interacts with GyrB as measured by a bacterial two-hybrid assay. E. coli strains derived from BTH101 harboring plasmids expressing the indicated fusion protein pairs were assayed for β-galactosidase activity indicative of protein-protein interactions. Experiments were performed in triplicate, and similar results were obtained from three independent experiments. V, control plasmid. (B) Interactions between GyrB and Fic-1 in vitro. GyrB-His6 was incubated with Affigels coated with Fic-1, Fic-1H135A, or BSA. After extensive washing, bound proteins eluted with SDS sample buffer were resolved by SDS-PAGE and detected by Coomassie brilliant blue staining. Ten percent of the GyrB used for the binding assay was loaded as reference. Data shown are representative of three experiments with similar results.

Fic-1 AMPylates GyrB on a tyrosine residue important for binding ATP

Because the Fic domain is capable of catalyzing diverse biochemical reactions (20), we attempted to determine whether GyrB is a target of Fic-1 by constructing an E. coli strain that coexpressed these two proteins tagged with glutathione S-transferase (GST) and His6, respectively. Purified GyrB-His6 protein was then separated by SDS–polyacrylamide gel electrophoresis (SDS-PAGE), excised, and digested with trypsin. The resulting peptides were analyzed by liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS). Spectra derived from MS/MS analysis of peptides were inspected by searching for peptides with potential change in molecular mass; a mass shift of 329 daltons corresponding to the addition of an AMP moiety was detected in the tryptic peptide -F104DDNSYK110- in GyrB coexpressed with wild-type Fic-1 but not with the Fic-1H135A mutant (Fig. 4A, upper panel). In addition, an intense peak of the diagnostic ion at 136.06 mass/charge ratio (m/z) (Fig. 4A, lower panel), corresponding to adenine derived from the breakdown of AMP, unambiguously indicated AMPylation. The unmodified fragment was almost undetectable in GyrB coexpressed with wild-type Fic-1, indicating that the modification was extensive (Fig. 4A, upper panel, middle). As a reference, the abundance of the tryptic peptide -G699LLEEDAFIER709- outside the modification site was very similar in these two samples (Fig. 4A, upper panel, right). Further analysis of the y and b series of the peptide backbone, more specifically, y1 and y2 fragments, allowed us to unambiguously assign the site of modification to the Tyr109 residue (Fig. 4A, lower panel).

(A) Identification of AMPylation of GyrB by MS. The upper panel is the extracted ion chromatograms of the unmodified and adenylylated versions of the peptide -F105DDNSYK110- derived from GyrB coexpressed with Fic-1 (left panel) or mutant Fic-1H135A (middle panel). The abundance of the randomly chosen peptide -G699LLEEDAFIER709- is almost identical in these two samples (right panel). The lower panel is the tandem mass spectrum of peptide -F105DDNSYK110-. The asterisks represent neutral losses of the adenosine moiety of the adenylylation modification. F, immonium ion of phenylalanine residue. (B) In vitro AMPylation of GyrB by Fic-1. After incubation at 35°C for 30 min, 32P-α-AMP–labeled signals and total proteins were detected by autoradiography for 30 sec (upper panel) or by staining (lower panel). (C) AMPylation of GyrB abolishes its ATPase activity. Samples from 30-min reactions were examined for ATP hydrolysis by measuring released phosphate (left panel). Induction of the SOS pathway by Fic-1 or Fic-1H135A was evaluated by the levels of LexA and RecA with ICDH as a loading control (middle panel). The band intensities of RecA (upper right) and LexA (lower right) were measured by LI-COR. All results are from three independent experiments. ***P < 0.0001.

To verify the results of the MS analysis, we analyzed Fic-1–mediated AMPylation of GyrB in vitro. Fic-1–His6, Fic-1H135A–His6, GyrB-His6, and GyrBY109F-His6 were purified, and the activity of Fic-1 and the mutant was evaluated using 32P-α-ATP. In the reaction containing wild-type Fic-1 and GyrB, robust 32P-AMP labeling of GyrB was detected, with an exposure time of less than 1 min (Fig. 4B, upper panel, third lane); self-modification of wild-type Fic-1 was also detectable (Fig. 4B, upper panel, second to fourth lanes). In reactions containing Fic-1H135A, no 32P-AMP-GyrB was detected (Fig. 4B, upper panel, fifth to seventh lanes), indicating a Fic motif–dependent activity. GyrB contains two distinct domains (35). We found that the N-terminal domain (residues 1 to 200), but not the C-terminal domain, of GyrB was modified by Fic-1 (fig. S6), indicating that this portion of the topoisomerase is sufficient to interact with the Fic-1. In agreement with the high degree of conservation with its E. coli counterpart, GyrB from P. fluorescens was modified by Fic-1 at Tyr111 (fig. S7), which corresponds to Tyr109 of the E. coli GyrB. The multiple modified GyrB products detected in these reactions may result from modification of partially degraded GyrB or the degradation of modified GyrB (fig. S7).

Weak but readily detectable signals in the position of GyrB were detected in the sample containing only wild-type Fic-1 (Fig. 4B, upper panel, second lane). Because this signal was also detected in experiments in which this sample was loaded far from the sample with strong modification (fig. S6, sixth and eighth lanes), we predicted that this represents AMPylated native GyrB that copurified with Fic-1. The lack of such signals in the reaction of wild-type Fic-1 with GyrBY109F may be due to the competitive effect of the excess GyrB mutant used in the reaction (Fig. 4B, upper panel, fourth lane). These results further validated GyrB as the target of Fic-1.

AMPylation by Fic-1 inactivates the ATPase activity of GyrB and induces the SOS response

GyrB is a subunit of DNA gyrase (DNA gyrase is also known as DNA topoisomerase II), the main activity of which is to introduce negative supercoiling to relieve the strain caused by helicase during DNA replication, in a process powered by ATP hydrolysis (36). In E. coli GyrB, Tyr109 is critical for binding ATP by forming a hydrogen bond with the N3 atom of the adenine ring (35). To examine the effects of Tyr109 AMPylation on the activity of GyrB, we determined the ATPase activity (37, 38) of GyrB that had been co-incubated with Fic-1. Whereas untreated GyrB or GyrB that had been co-incubated with Fic-1H135A exhibited readily detectable ATP hydrolysis (Fig. 4C, left panel), neither Fic-1 nor its H135 mutant hydrolyzed ATP, nor did GyrB with a mutation in Tyr109 (Fig. 4C, left panel). As expected, GyrB that had been co-incubated with Fic-1 and ATP displayed significantly lower ATPase activity (Fig. 4C, left panel). Thus, AMPylation of GyrB by Fic-1 abolished the ATPase activity of GyrB.

Inhibition of GyrB activity causes DNA replication arrest, which results in the exposure of single-strand DNA and the subsequent formation of RecA filaments (39, 40). This facilitates the autocatalytic cleavage of the LexA repressor and the subsequent induction of the SOS response (41). Consistent with this pattern, the amount of RecA increased significantly in cells expressing Fic-1, but not in those expressing the Fic-1H135A mutant, whereas the amount of LexA, the SOS repressor, decreased in a Fic-1–dependent manner (Fig. 4C, right and lower panels). Novobiocin, which selectively inactivates GyrB, served as a positive control for induction of the SOS response.

Self-AMPylation of Fic-1 is important for its activity on GyrB

We consistently observed self-AMPylation of Fic-1, particularly in reactions lacking its substrate GyrB (Fig. 4B, upper panel, second lane). To explore the potential role of self-modification on Fic-1 function, we identified the modified site by MS analysis. Tyr5 was AMPylated in Fic-1 incubated with ATP (Fig. 5A). Substitution of Tyr5 with an Ala residue resulted in no detectable self-AMPylation (Fig. 5B and fig. S8). The Fic-1Y5A mutant failed to AMPylate GyrB (Fig. 5B). In agreement with the loss of AMPylator activity toward GyrB, Fic-1Y5A no longer affected the yield of plasmid DNA in E. coli (Fig. 5C). Together, these results indicate that self-AMPylation is critical for this function of Fic-1.

(A) Self-modification by Fic-1 occurs on Tyr5. Fic-1 co-incubated with ATP was analyzed by MS after trypsin digestion. The upper panel is the extracted ion chromatograms of the unmodified and adenylylated versions of the tryptic peptide containing Tyr5. The abundance of the randomly chosen peptide -N179GVMEPMEQVFEK191- is almost identical in these two samples (right panel). The lower panel is the tandem mass spectrum of the AMPylated peptide -Y5GVGEDAYCYPGSTVLR21-. CAM, carbamidomethylation. (B) A mutation in Tyr5 abolishes self-AMPylation of Fic-1. Reactions containing the indicated proteins were incubated at 35°C for 30 min, AMPylation was detected by autoradiography for 2 min (upper panel), and total proteins were visualized by staining (lower panel). WT, wild type. (C) The Y5A mutation abolishes the ability of Fic-1 to inhibit plasmid DNA yield. DNA of plasmids carrying fic-1 or the indicated mutants isolated from identical amounts of E. coli cells was digested with restriction enzymes Bam HI and Sal I and was separated using agarose gels (upper panel). An identical set of samples was processed for immunoblotting to detect the levels of Fic-1 (lower panel). The images in (B) and (C) are representative of three independent experiments with similar results.

The activity of Fic-1 is regulated by a specific inhibitor

Analysis of the DNA sequence upstream of the fic-1 open reading frame identified a gene potentially encoding a protein of 56 residues (fig. S1). These two genes overlap for one nucleotide. Sequence analysis revealed that the protein encoded by this gene harbors an -S24LRLEG29- motif, which is found in α-inhibitors of Fic proteins (32). When expressed in E. coli, this gene rescued the inhibition of plasmid DNA replication conferred by Fic-1 (fig. S9A), suggesting that the protein functioned as an α-inhibitor. We designated this gene as antF (anti–Fic-1). AntF interacted with Fic-1 in a bacterial two-hybrid assay; Cya fusions of AntF and Fic-1 drove the expression of the LacZ reporter to levels comparable to that of leucine zipper proteins used as positive controls (34) (fig. S9B). Similar results were obtained with a reciprocal fusion orientation (fig. S9B). In agreement with the genetic results, inclusion of recombinant AntF in the AMPylation reaction with Fic-1 reduced the modification of GyrB in a dose-dependent manner (Fig. 6, A and B). Even a 1:27 molar ratio of Fic-1 to AntF1 significantly inhibited the AMPylation activity, and a 1:3 ratio completely abolished activity.

(A) Dose-dependent inhibition of Fic-1 by AntF. The indicated amounts of His6-SUMO-AntF were added to a series of identical reactions containing Fic-1. After 30 min of incubation, equal amounts of a mixture containing GyrB and 32P-α-ATP were added, and the reactions were allowed to proceed for 30 min at 35°C. 32P-α-GyrB and total proteins were detected by autoradiography (upper panel) and Coomassie blue staining (lower panel). (B) Quantification of 32P-α-GyrB signals. The strength of autoradiography signals from three independent experiments done under the same conditions was measured and analyzed by ImageJ. **P < 0.01.

We attempted to examine the potential phenotypes of mutants lacking the α-inhibitor. Despite repeated attempts, we could not delete the inhibitor-encoding gene without compromising the expression of fic-1 in P. fluorescens strain 2P24 (Fig. 7A), suggesting that this bacterium cannot tolerate uncontrolled activity of Fic-1 expressed from its cognate promoter. These results are consistent with the observation that ectopically expressed Fic-1 in P. fluorescens was lower than the mutant, although the induction of filamentation and growth arrest were apparent (Fig. 2B and fig. S5C).

Fig. 7Expression of Fic-1 in P. fluorescens strains and induction of cell filamentation by Fic-1 in E. coli defective in sulA and recA.

(A) Bacterial strains were grown in LB to an optical density at 600 nm (OD600) of 2.4, and protein samples prepared from 300-μl cultures were resolved by SDS-PAGE. Fic-1 was probed with a specific antibody. A protein nonspecifically recognized by the antibody at about 30 kD served as a loading control. Note that Fic-1 is detectable only in the WT. (B) Morphology of cells expressing Fic-1. Cells of transformants grown for 16 hours on LB agar resuspended in PBS were fixed and stained with Hoechst. Images acquired with a fluorescence microscope were pseudocolored with IPLab software. Images shown are representative of three experiments with similar results. (C) Distribution of cells with different lengths. The length of 500 cells was measured from each of three samples, and their distribution was plotted. Data shown are representative from three independent experiments.

When the SOS response is blocked, Fic-1 triggers cell elongation

Activation of the SOS pathway involves the formation of RecA filaments (40, 42), the auto-cleavage of the SOS repressor LexA (43), and the induction of sulA, which binds FtsZ, thus blocking the completion of cell division and leading to the formation of filamentous cells (12). To examine whether cell filamentation induced by Fic-1 required the SOS pathway, we introduced constructs harboring arabinose-inducible Fic-1 into an E. coli strain defective in recA and sulA. Expression of Fic-1, but not Fic-1H135A, caused morphological changes. In the strain expressing Fic-1, cells longer than 10 μm were readily detectable and more than 50% of cells were longer than 2.5 μm, whereas most of the cells expressing the Fic-1H135A mutant displayed lengths between 1 and 1.75 μm, which were similar to those in the vector control (Fig. 7, B and C). Whereas some of the cells expressing Fic-1 were filamentous, most cells displayed an elongated morphology with a single nucleoid (Fig. 7, B and C). These results suggested that in cells defective in the SOS pathway, inhibition of DNA replication by Fic-1 did not block cell elongation.

Despite repeated attempts, we were unable to construct a derivative of the ∆sulA strain of P. fluorescens expressing Fic-1 (no transformants were obtained in experiments aiming at introducing the plasmid that directs the expression from the arabinose-inducible promoter into the ∆sulA strain). This observation further indicates that P. fluorescens is more sensitive to the Fic protein than E. coli is.

DISCUSSION

Fic proteins are widespread across all domains of life, with the largest number present in taxonomically diverse bacteria (19). Despite the fact that high-resolution structures of a number of bacterial Fic proteins are available, our understanding of the biological activities of this family comes primarily from bacterial virulence factors (24) and the Doc protein of phage P1 that targets host protein translation as a Fic domain–mediated kinase (24, 44). Our results establish that a housekeeping Fic protein AMPylates GyrB, a subunit of the bacterial type II DNA topoisomerase, thus interfering with DNA replication. The modification by Fic-1 abolishes the activity of GyrB and induces filamentation.

Several lines of evidence show that Fic-1 specifically targets GyrB. First, Fic-1 but neither Fic-2 nor Fic-3 from the same bacterium interferes with bacterial growth or plasmid replication. Second, Fic-1 is able to induce cell filamentation, a phenotype often associated with the inhibition of GyrB activity (38, 45). Third, GyrB is the only protein that interacts with Fic-1 among the 77 DNA replication proteins tested. In addition, native GyrB was copurified with Fic-1–His6 (Fig. 4B and fig. S6), consistent with the sequence of its Fic domain (H135PFREGNGR143), which predicts an adenylyl transferase activity (24, 44). Fic-1 targets the GyrB subunit of type II topoisomerase by adding an AMP moiety to the highly conserved tyrosine residue Tyr109 essential for binding ATP. Antibiotics of the coumarin and cyclothialidine families inhibit the ATPase activity of DNA gyrase by blocking the binding of ATP to GyrB (46). Our results are consistent with a recent report showing the targeting of DNA gyrase and topoisomerase IV by several Fic proteins (47). It appears that interference with the ATP binding of GyrB is a common mechanism for inhibiting its activity. This is most likely because ATPase activity is essential for GyrB function, and inhibition of ATP binding probably is the most effective means to block its activity.

The presence of a specific and effective inhibitor of Fic-1 indicates tight regulation of its activity. The fact that we were unable to delete the inhibitor gene in strain 2P24 further indicates the importance of such regulation, which is consistent with the observation that ectopic expression of Fic-1 inhibits bacterial growth. Apparently, the inhibitory effects of AntF need to be eliminated when cell filamentation or cell division arrest has become necessary. We propose that such elimination can be achieved by at least two mechanisms, both involved in the induction of a signaling cascade in response to appropriate environmental cues (Fig. 8). In the first scenario, the signaling events induce the expression of a protease that degrades AntF, whereas in the second scenario, the titration of the inhibitor is mediated by a protein to which it has a higher affinity than to Fic-1 (Fig. 8).

Fig. 8A model of Fic-1–mediated induction of bacterial cell filamentation and its regulation.

Fic-1 and the α-inhibitor AntF form a dynamic complex under normal conditions. Inducing signals from the environment activate a cascade that leads to the production of a sequestering protein (Seq) that competes for AntF or the activation of a protease that degrades the α-inhibitor. Freed or activated Fic-1 then inactivates GyrB by AMPylation, leading to the induction of SulA and the formation of filamentous cell. Alternatively, AMPylated GyrB may induce cell filamentation through an SOS response–independent pathway (dashed arrows).

Many bacteria, including E. coli and Pseudomonas spp., become filamentous under stress conditions such as DNA damage caused by factors like ultraviolet (UV) radiation or partial inhibition of cell wall biosynthesis by antibiotics (1). UPEC forms filaments only in immune competent hosts (3). Filamentous bacteria are more resistant to engulfment by phagocytes and other predators (1). In parallel, inhibition of DNA replication will most likely lead to slower metabolic rates, making bacteria resistant to some antibiotics and to certain detrimental environmental conditions. Regulation of cell morphology by targeting DNA replication has been described for the SocA-SocB TA module in Caulobacter crescentus (48). Similarly, Fic-induced filamentation has been reported, although the mechanism is unknown (32). The identification of GyrB as a target of Fic-1 demonstrates the function of at least a subset of widely distributed Fic proteins in the regulation of bacterial cell division. Our results suggest that in response to Fic-1 activity, a yet unidentified SOS-independent pathway contributes to cell morphology changes (Fig. 8). SOS-independent induction of cell filamentation has been documented in C. crescentus (49). It will be interesting to determine whether the SOS-independent pathways involved in the regulation of cell morphology are conserved among bacterial species.

Self-modification occurs in many Fic proteins, which has been used to monitor their biochemical activity when a substrate is not available (32). Our finding that the Fic-1Y5A mutant defective in self-AMPylation had almost completely lost its activity against GyrB implies an important role of self-AMPylation in its function. In its complex with the α-inhibitor VbhA, the N-terminal end of VbhT (its fourth residue is a Tyr) does not seem to participate in the formation of the catalytic pocket (32). It is possible that in the absence of the inhibitor, this Tyr assumes an important role in the overall conformation of these Fics or in the formation of the catalytic pocket. Determining the role of this Tyr4 will likely reveal whether this residue is generally important for the activity this groups of Fic proteins.

Pressure from the immune system and metabolic fluctuations often result in persister cells that are resistant to immune killings as well as to antibiotics (46). In E. coli, TA modules are involved in the formation of persisters (50). FicTAs may participate in the establishment of persisters or in the regulation of the transition between the bacillary and filamentous forms during infection. One future research avenue is the identification of the pathway that parallels the SOS cascade in regulating bacterial morphology. Similarly, it will be of great interest to determine whether any of these stresses, such as antibiotic, UV radiation, and immune suppression, cause Fic-1 activation, and if so, the mechanism underlying such derepression.

MATERIALS AND METHODS

Bacterial strains, plasmids, and media

Bacteria and plasmids used in this study are listed in table S2. P. fluorescens strain 2P24 and its derivatives were grown in LB medium or Agrobacterium mannitol (ABM) minimal medium (51) at 28°C. E. coli strains were incubated at 37°C with the exception of strain BTH101 used for bacterial two-hybrid assays (34), which was cultured at 30°C. For plasmid selection, antibiotics were supplemented in media at the following concentrations: ampicillin, 50 μg/ml; kanamycin, 50 μg/ml; gentamicin, 10 μg/ml; chloramphenicol, 20 μg/ml; and tetracycline, 20 μg/ml. When needed, 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-gal) was used at 20 μg/ml. To induce SOS in E. coli, novobiocin was added to a final concentration of 25 μg/ml to bacterial cultures established by 10-fold dilution of overnight cultures. The cells were incubated at 37°C in a shaker (200 rpm/m) for 6 hours before processing for SDS-PAGE and immunoblotting.

Bacterial strain construction

In-frame deletion mutants of P. fluorescens strain 2P24—2P24∆fic-1, 2P24∆fic-1∆antF, 2P24∆αantF, and 2P24∆sulA—were constructed by a two-step gene replacement procedure as described previously (33). The success of deletion was verified by colony polymerase chain reaction (PCR) with primers flanking the genes of interest.

Site-directed mutagenesis

Mutations in specific site of the genes of interest were introduced by QuikChange II (Agilent Technologies) with the PfuUltra High-Fidelity DNA polymerase and overlapping primer pairs bearing the desired mutations. The specificity of the mutations was verified by double-stranded DNA sequencing.

Evaluation of plasmid DNA yield in E. coli

The coding regions of fic-1, fic-2, and fic-3 amplified by PCR with appropriate primers (table S2) from genomic DNA of P. fluorescens strain 2P24 (33) were digested with appropriate restriction enzymes. Bam HI and Sal I were used for fic-1 and Hind III and Xba I were used for fic-2 and fic-3 for inserting into pHSG399 (52) to give pCL001, pCL002, and pCL003, respectively. The expression of the fic genes in these plasmids is driven by the lac promoter. The histidine residue in the Fic motif of fic-1 was mutated to an alanine by site-directed mutagenesis to give pCL004 (Fic-1H135A). To test the effects of the predicted α-inhibitor, the region containing both the inhibitor and fic-1 genes was inserted pHSG399 to make pCL005. These plasmids were individually transformed into E. coli strain DH5α. Bacteria were grown overnight at 37°C in 3 ml of LB broth containing chloramphenicol (20 μg/ml). On the basis of the cell density (OD600), two identical samples with the same concentration of cells were withdrawn. One set of samples was subjected to plasmid DNA isolation with the Zyppy Plasmid Miniprep kit (Zymo Research), eluting twice, each with 50 μl of TE buffer (pH 8.0). DNA samples (10 μl) were digested with Bam HI and Sal I or Hind III and Xba I, the two restriction enzymes used for subcloning of the genes. The DNA fragments were separated by agarose gel electrophoresis, and the gels were stained with ethidium bromide (1 μg/ml) for 15 min. Images were acquired with a Bio-Rad gel documentation system. The second sample set was used for immunoblotting to detect ICDH as a loading control.

To tightly regulate gene expression, fic-1 and the fic-1H135A mutants were subcloned into pBAD22 containing the arabinose-regulatable PBAD promoter (53). To express wild-type fic-1 in P. fluorescens, a tightly regulated plasmid was developed by inserting the PBAD promoter into pBBR1MCS-2 (54) to give pCL008, which was used to express fic-1 and the fic-1H135A mutant genes. The relevant plasmids were introduced into P. fluorescens strains PM933, PM937, and PM938 (table S2). Induction of expression from the PBAD promoter was performed in ABM minimal medium with 0.2% arabinose.

Bacterial two-hybrid and β-galactosidase activity assays

The Cya-based bacterial two-hybrid system (34) was used to examine interactions of proteins. Briefly, Fic-1 was fused to the T25 fragment of the Cya in pKT25 (34), and proteins known to be involved in DNA replication (table S1) were individually fused to the T18 fragment of adenylyl cyclase in pUT18C (34). Plasmid pairs were introduced into strain BTH101 (34), and the interactions were first evaluated on LB media containing X-gal. Strains exhibiting positive interactions indicated by the hydrolysis of X-gal to form blue colonies were retained for further analysis by quantitative measurement of β-galactosidase activity in liquid cultures.

To measure β-galactosidase activity, overnight bacterial cultures were diluted into fresh media at a ratio of 1:20. Cells (100 μl) from saturated cultures were collected for enzymatic assays with ONPG (o-nitrophenol-β-d-galactoside) following the standard protocol (55). All assays were performed in triplicate, and the activity was expressed in Miller units.

Purification of recombinant proteins

Unless otherwise stated, E. coli strain BL21(DE3) was used to express recombinant protein. To express Fic-1 or Fic-1H135A, the coding regions amplified by PCR (primer information in table S2) were digested with Nde I and Sal I before inserting into similarly digested pET22b(+) to give pET22b–Fic-1 and pET22b–Fic-1H135A. To purify potential substrates of Fic-1 and other proteins such as AntF, their coding regions amplified by PCR were inserted into appropriately digested pET-SUMO that allows the production of His6-SUMO–tagged proteins (56). The integrity of each gene in the expression vector was verified by double-stranded sequencing.

For protein production, strains derived from BL21(DE3) harboring the appropriate expression plasmids were cultured overnight in 5 ml of LB broth at 37°C, which were then transferred to 500 ml of broth and grown at 37°C to an OD600 of 0.6 to 0.8. After adding IPTG to a final concentration of 250 μM, incubation was continued at 18°C in a shaker (200 rpm) for 16 hours. The cells were harvested by centrifugation at 6000g for 10 min at 4°C. Unless otherwise indicated, the pellets were suspended in PBS containing 20 mM imidazole and protease inhibitors (1 μM phenylmethylsulfonyl fluoride and 5 μM benzamidine) before lysis by sonication. Cell debris and unbroken cells were removed by centrifugation twice at 12,000g for 15 min at 4°C. The cleared lysates were incubated with 2 ml of Ni-NTA beads (Qiagen) for 2 hours at 4°C. Beads with bound proteins were loaded onto a column. After washing the column with 3 × column volume of PBS buffer containing 20 mM imidazole, protein was eluted with PBS containing 250 mM imidazole. Proteins were dialyzed in PBS buffer with the addition of 20% (v/v) glycerol and 0.5 mM dithiothreitol (DTT). When needed, the SUMO-specific protease Ulp1 (56) was used to cleave the His6-SUMO tag from the recombinant proteins by incubating at 30°C for 3 hours. Proteins in solution were dialyzed in appropriate buffers.

To purify recombinant GyrB and its mutants, the coding regions were amplified by PCR with primers (table S2), which introduced six histidine residues at the carboxyl termini of the proteins. The products were digested with Bam HI and Xho I and inserted into Bam HI/Sal I digested pGEX-6P1 to give pCL009. Plasmids were introduced E. coli strain XL1Blue and cells induced to express the proteins were prepared as described above. For purification, cells were lysed in tris buffer [50 mM tris-HCl (pH 7.4), 150 mM NaCl], and the same buffer containing 20 mM imidazole was used to wash the column loaded with the protein-bound beads. The target protein was eluted with tris buffer (50 mM tris-HCl, pH 7.4) containing 200 mM imidazole. After dialysis against tris buffer [50 mM tris-HCl (pH 7.4), 0.5 mM DTT, and 20% (v/v) glycerol], the GST tag was removed with the PreScission protease by incubation overnight at 4°C. The GST tag and the protease were removed with Glutathione Sepharose 4 Fast Flow beads (GE Healthcare). The purity of all proteins was 95% or higher as evaluated by Coomassie brilliant blue staining after SDS-PAGE.

Bacterial growth analysis

To evaluate the effects of Fic-1 on bacterial growth, E. coli strain BL21(DE3) was transformed with pHisSUMO, pHisSUMO–Fic-1, or pHisSUMO–Fic-1H135A. Transformants were selected on LB plates containing kanamycin. Growth was documented by acquiring the images of the colonies appearing on the same plate after 16 hours of incubation at 37°C. To evaluate the growth in liquid medium, transformants patched onto LB plates were grown overnight at 37°C; the cells on patches were then collected and washed twice with PBS. The densities of the cell suspensions were measured by optical density after appropriate dilutions. Cultures of identical density were established in 150 ml of LB broth. After incubation at 37°C for 30 min, each culture was split into three subcultures and incubated in a shaker at 200 rpm. The growth of the cells was monitored by measuring OD600 at 2-hour intervals. The averages of the readings from the three independent cultures were plotted against the incubation time.

For P. fluorescens, derivatives of strain 2P24 harboring the appropriate plasmids were grown to saturation for 14 hours in LB medium containing 0.2% glucose. The cultures were 1:100 in ABM medium (51) containing 0.2% arabinose, and growth was monitored by measuring OD600 at the indicated time points.

Coexpression of GyrB and Fic-1

Plasmid pCL001 (pHSG399::fic-1) (table S2) was cotransformed with pGyrB-His6 (pCL009) into strain XL1Blue and the resulting strain was grown in 5 ml of LB broth to saturation. A 200-ml culture established by a 1:20 dilution of the overnight culture was grown to an OD600 of 0.6 to 0.8, IPTG was added to 100 μM, and the culture was further incubated at 16°C in a shaker for 14 hours to induce expression of the proteins. Cells were used to purify GyrB-His6 as described above.

Mass spectrometry

Purified proteins were resolved by SDS-PAGE and the bands corresponding to GyrB or Fic-1 were excised from acrylamide gels and digested with trypsin (57). The resulting peptides were analyzed by LC MS/MS on an Ekspert nanoLC 400 (Eksigent) connected to a 5600 TripleTOF mass spectrometer (AB Sciex). Peptides were loaded into a C18 trap column (200 μm × 0.5 mm, ChromXP C18-CL, 3 μm, 120 Å, Eksigent) and the separation was carried out in a capillary C18 column (75 μm × 15 cm, ChromXP C18-CL, 3 μm, 120 Å) at 200 nl/min using the following gradient: 5% solvent B [solvent A: 0.1% fatty acid (FA) and solvent B: 80% acetonitrile/0.1% FA] for 1 min, 5 to 35% solvent B for 60 min, 35 to 80% solvent B for 1 min, 80% solvent B for 6 min, 80 to 85% solvent B for 1 min, and hold at 5% for 11 min. Full MS spectra of eluting peptides were collected in the range of 400 to 2000 m/z, and the 10 most intense parent ions were submitted to fragmentation for 250 ms using rolling collision energy. The spectra were analyzed manually by de novo sequencing.

In vitro binding between Fic-1 and GyrB

Water-activated Affigel 15 (Bio-Rad) beads were coated with 50 μg of Fic-1–His6 or Fic-1H135A–His6 at 4°C for 14 hours. Beads coated with the same amount of BSA were also prepared. After washing with three bed volumes of PBS containing 1 mM DTT and 1% Triton X-100, the beads were blocked with 20 mM tris (pH 8.0) at 4°C for 2 hours. Washed beads were split into identical samples and 40 μg of GyrB-His6 was added into each reaction. Ten percent of the samples were withdrawn as input controls, and the bead slurry was incubated at 4°C on a rotary shaker for 4 hours. Unbound proteins were removed by washing the beads with five bed volumes of PBS, and bound proteins were extracted with SDS loading buffer. Proteins separated by SDS-PAGE were detected with Coomassie brilliant blue staining.

In vitro AMPylation assay

The AMPylation assay was performed as described previously (58). Briefly, 20-μl reactions containing 1.5 μg of Fic-1–His6, 10 μg of GyrB-His6, and 5 μCi of 32P-α-ATP (PerkinElmer) in a reaction buffer [25 mM tris-HCl (pH 7.5), 50 mM NaCl, 3 mM MgCl2, 0.5 mM EDTA] were allowed to proceed for 30 min at 35°C. The reaction was terminated with 5 μl of 5× Laemmli buffer. After denaturing by boiling for 5 min, samples were separated by SDS-PAGE. The gels were first stained with Coomassie brilliant blue to evaluate protein levels in the reactions and were then dried before x-ray film autoradiography to detect AMP-α–labeled molecules.

For dose-dependent AMPylation of GyrB by Fic-1, a 120-μl master reaction containing 60 μg of GyrB-His6 was established in the AMPylation buffer. Six subreactions were then established by aliquoting the master reaction into one reaction of 30 μl, four reactions of 20 μl, and one reaction of 10 μl. Fic-1–His6 (6.75 μg) was added to the 30-μl reaction, and the 10-μl mixed sample was added to one of the 20-μl reactions to perform a threefold serial dilution. One of the 20-μl reactions was set as a Fic-1–free control. The reaction was started by adding 5 μCi of 32P-α-ATP into each test tube. After incubation at 35°C for 30 min, reactions were terminated, and the samples were subjected to SDS-PAGE and detection by Coomassie brilliant blue staining as described above.

To test the effects of the AntF on the activity of Fic-1 on GyrB, identical subreactions were prepared from a 120-μl master reaction containing 9 μg of Fic-1–His6, into which the indicated amounts of His6-SUMO-AntF were then added, and the tubes were incubated at room temperature (RT) for 30 min. A reaction that did not receive His6-SUMO-AntF was used as a control. A solution containing 32P-α-ATP and 10 μg of GyrB-His6 was aliquoted into these reactions. After incubation at 35°C for 30 min, samples resolved by SDS-PAGE were analyzed for protein levels and for 32P-α-ATP–labeled signals as described above.

Fluorescence microscopy analysis

Bacterial cells from transformants appearing on selective medium were suspended in PBS and fixed with 4% paraformaldehyde following an established protocol (59). Tween 20 was added to 0.5‰ diluted samples; 10 μl from each sample was dropped onto coverslips and dried over a flame, and nonadherent cells were removed with PBS. To stain DNA, the coverslips were placed in 50 μl of Hoechst (10 μg/ml) at RT for 30 min. After washing three times with PBS, the coverslips were mounted on glass slides with antifade reagent (Vector Laboratories). Samples were inspected with an Olympus IX-81 fluorescence microscope, and images were acquired using a charge-coupled device camera with identical digital imaging parameters (objectives, exposure duration, contrast ratios, etc.). The images were similarly processed using the IPLab software package (BD Biosciences).

ATPase assay

To measure ATPase activity of GyrB, all proteins were purified in tris buffer. The ATPase/GTPase Activity Assay Kit (MAK113, Sigma) was used in 96-well plates at RT (22°C). GyrB (5 μg) or GyrBY109F (5 μg) was preincubated with the indicated amounts of Fic-1 or Fic-1H135A in 20-μl assay buffer (40 mM tris, 80 mM NaCl, 8 mM MgAc2, and 1 mM EDTA, pH 7.5), and the final volume was adjusted to 30 μl with deionized H2O. Similar reactions without any protein or only one of the proteins being tested were set up as controls. To test the dose-dependent activity of Fic-1, 5 μg of GyrB was preincubated with increasing amounts of Fic-1. The reactions were initiated by adding 10 μl of 4 mM ATP and were allowed to proceed for 30 min before being terminated with 200 μl of malachite green. After 30 min at RT, the intensity of the signal was measured by determining the absorbance at 620 nm. The concentration (micromolar) of free phosphate in the reactions was calculated from a standard curve using phosphate standard supplied in the kit.

Antibodies and immunoblotting

Fic-1–specific antibodies were generated with Fic-1–His6 by Pocono Rabbit Farm and Laboratory following standard protocols. Antibodies were affinity-purified using Affigel 15 (Bio-Rad) coated with Fic-1–His6 following a standard protocol (29). The antibody was used at 1:30,000 for immunoblotting. Antibody against α-ICDH (59) was used at 10,000. Antibodies against RecA and LexA were purchased from Santa Cruz Biotechnology and Abcam, respectively, and were used at 1:3000 and 1:5000.

For immunoblotting, proteins resolved by SDS-PAGE were transferred onto nitrocellulose membranes. The membranes were blocked with 5% (w/v) nonfat dried milk in PBST (PBS + 0.2% Tween 20) buffer. After being washed three times with the buffer, the membranes were incubated with the primary antibody at RT for 1 hour. Similarly washed membranes were incubated with an appropriate IRDye infrared secondary antibody (LI-COR Biosciences), and the signals were detected; if necessary, the intensity of the bands was quantitated by using the LI-COR Odyssey imaging system.

Data quantitation and statistical analyses

The images captured from DNA agarose gels by a Bio-Rad Universal Hood II Gel Doc (Bio-Rad Laboratories) were quantified by using the Quantity One 4.6.9 software (Bio-Rad Laboratories). ImageJ was used to analyze autoradiograph films in AMPylation assays after scanning by Artix Scan M1 (Microtek). Replications of these experiments were scanned and analyzed with the same parameters. Immunoblots were scanned and quantified using the Odyssey 3.0 (LI-COR Biosciences). Student’s t test was used to compare the mean levels between two groups, each with at least three independent samples.

Acknowledgments: We thank A. Aronson for critical reading of the manuscript, and L. Csonka and the Coli Genetic Stock Center at Yale University for bacterial strains. Funding: This work was supported by NIH grants R56AI103168 and K02AI085403 to Z.-Q.L., National Natural Science Foundation of China grants (31272082, 31572045) to L.-Q.Z., and the 111 Project B13006 to Y.-L. Peng (China Agricultural University). C.L. was supported by a fellowship from the China Scholarship Council. Author contributions: L.-Q.Z. and Z.-Q.L. designed the study and supervised the work. C.L. performed the genetic and biochemical experiments. E.S.N. performed the MS experiments and analyzed the data. C.L. and Z.-Q.L. wrote the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: The MS data are available at ProteomeXchange repository (www.proteomexchange.org/) under accession numbers PXD003077 and PXD003076. GenBank accession numbers for the genes described in this study: antF, KT020754; fic-1, KT020755; fic-2, KT020756; fic-3, KT020757; gyrB, KT020758.